Alkylation of nucleobases by 2-chloro-N,N-diethylethanamine hydrochloride (CDEAH) sensitizes PARP1-deficient tumors

Abstract Targeting BRCA1- and BRCA2-deficient tumors through synthetic lethality using poly(ADP-ribose) polymerase inhibitors (PARPi) has emerged as a successful strategy for cancer therapy. PARPi monotherapy has shown excellent efficacy and safety profiles in clinical practice but is limited by the need for tumor genome mutations in BRCA or other homologous recombination genes as well as the rapid emergence of resistance. In this study, we identified 2-chloro-N,N-diethylethanamine hydrochloride (CDEAH) as a small molecule that selectively kills PARP1- and xeroderma pigmentosum A-deficient cells. CDEAH is a monofunctional alkylating agent that preferentially alkylates guanine nucleobases, forming DNA adducts that can be removed from DNA by either a PARP1-dependent base excision repair or nucleotide excision repair. Treatment of PARP1-deficient cells leads to the formation of strand breaks, an accumulation of cells in S phase and activation of the DNA damage response. Furthermore, CDEAH selectively inhibits PARP1-deficient xenograft tumor growth compared to isogenic PARP1-proficient tumors. Collectively, we report the discovery of an alkylating agent inducing DNA damage that requires PARP1 activity for repair and acts synergistically with PARPi.


INTRODUCTION
Pol y(ADP-ribose) pol ymerase 1 (PARP1) promotes DNA repair by binding to DNA breaks and by attaching ADPribose polymers to itself and a number of other proteins to regulate DNA repair. PARP1 has been shown to have a role in man y DNA pathwa ys and has a special role in base excision repair (BER) ( 1 , 2 ). In addition, PARP1 has been shown to have a role in r egulating r eplication fork speed ( 3 , 4 ), protecting stalled replication forks ( 5 , 6 ) and pre v enting the formation of gaps formed on the lagging strand by incomplete Okazaki fragment synthesis in lagging strand DNA synthesis ( 3 , 5-8 ). Ther efor e, PARP1 dysfunction or inhibition in homologous recombination (HR)deficient cells leads to the accumulation of replication gaps in S phase, and the exposed lagging strand gaps become toxic to cells ( 7 ). Thus, inhibiting PARP1 is synthetic lethal to cells with defects in HR genes ( 9 , 10 ). Such synthetic lethality, referring to the cell-lethal effects upon the inactivation of two genetically distinct pathways, is a useful approach to selecti v ely kill cells with defects in a DNA repair pathway (11)(12)(13). In particular, synthetic lethality provides a conceptual frame wor k for the de v elopment of drugs that are selecti v ely toxic in specific genetic backgrounds associated with tumors. In addition to PARP1 inhibition, prominent recent examples are the use of PolQ inhibitors to inhibit alternati v e end joining pathways to target tumors carrying a mutation in the BRCA1 or BRCA2 ( 14 , 15 ) or inhibition of the Werner protein in tumors with mismatch repair (MMR) deficiencies ( 16 ).
Currently, certain types of breast (e.g., high-grade serous ovarian cancer or triple-negati v e breast cancer), ovarian, pancrea tic or prosta te cancers carrying a BRCA1 or BRCA2 mutation can be treated with PARP inhibitors (PARPi) as a first-line treatment ( 17 ). Although PARPi monotherapy has shown promising efficacy and safety profiles in clinical practice, its major limitations are the need for specific alterations in HR genes in tumors and the rapid emergence of resistance (18)(19)(20). Many tumors that initially respond to PARPi treatment e v entually recur through compensatory muta tions tha t r estor e HR acti vity or stimulate the acti vity of alternati v e repair pathways ( 11 ). To overcome these issues in clinical practice, various combinatorial treatments of PARPi with drugs targeting other pathways are currently being tested ( 11 , 17 ).
A TAD5 (A TPase family AAA domain-containing protein 5) is a human protein encoded by the ATAD5 gene that belongs to the AAA+ ATPase family. Its main function is to ensur e DNA r eplication and maintain genomic stability by r egulating DNA r eplication initiation and elongation, responding to DNA damage and stabilizing stalled replication forks (21)(22)(23)(24)(25)(26)(27)(28). ATAD5 d ysregula tion or muta tions have been linked to various types of cancer (29)(30)(31). ATAD5 is also a useful biomarker for detecting genotoxic compounds, as its protein le v els increase after DNA damage ( 32 ). To identify small molecules eliciting DNA replication stresses, a HEK293T cell line stably expressing the luciferase-tagged A TAD5 (A TAD5-luc cell), which measures the luciferase activity as a readout to measure the le v el of ATAD5 expression in response to DNA damage or replication stress ( 33 ), was used for the identification of small molecules inducing DNA replication stresses. We reasoned that through a screen for small molecules that cause r eplication str ess combined with an analysis of the pathway(s) inhibited by any of the hits, we would be likely to identify molecules that act synergistically with PARPi. Using this assay, we screened a 344,385 small molecule library (the National Institute of Health's Molecular Libraries Probe Production Centers Network) and identified 289 small molecules that activated expression of the ATAD5 reporter gene. Among the positi v e hits, we have already characterized the small molecule baicalein, which can selecti v ely kill MMR-deficient tumors through its pr efer ential interaction with mismatched DNA and the MSH2-MSH6 complex for activation of the ATM-CHK2 pathway ( 34 ).
To begin to identify the molecular mechanisms by which other identified small molecules cause DNA replication stress and DNA damage, we treated various cell lines with mutations in DNA repair genes to identify whether any of them caused synthetic lethality. In this screen, we identified 2-chloro-N , N -diethylethanamine hydrochloride (CDEAH), monofunctional or half-nitrogen mustard, as a small molecule that selecti v ely kills PARP1 -and xeroderma pigmentosum A ( XPA )-deficient cells in cell culture and x enograft models. CDEAH pr efer entially alkylates guanine residues in DNA, forming adducts that can be removed by either BER or nucleotide excision repair (NER). As the intermediate in BER, an abasic site is bound by PARP. Collecti v ely, we report a potential synergistic treatment option of PARPi by enhancing DNA damage that depends on PARP1-dependent BER mechanisms.

A T AD5-luciferase assay
HEK293T A TAD5-L UC cells ( 33 ) were plated at a density of 15,000 cells per well in a 96-well white, assay plate (Costar). After 24 h, cells were treated with 5-FUrd or CDEAH and incubated for an additional 24 h. Luciferase activity was measured by adding ONE-Glo luciferase reagent (Promega) to each well and measuring luminescence intensity with a Synergy NEO2 Hybrid Multi-Mode Reader (BioTek).

Immunoblot analysis
Whole-cell extracts were isolated and imm unoblot anal ysis was performed as previously described ( 24 ) with slight modifications. Briefly, w hole-cell extracts were isolated by incubating cells on ice with Benzonase ® nuclease (250 units / l, Enzynomics) in RIPA buffer [50 mM Tris-HCl (pH 8.0), 150 mM NaCl, 5 mM EDTA, 1% Triton X-100 ™, 0.1% sodium dodecyl sulfate, 0.5% sodium deoxycholate, Halt ™ Protease & Phosphatase Single-Use Inhibitor Cocktail] for 40 min, followed by sonication and centrifugation. For imm unoblot anal ysis, proteins wer e r esolved by sodium dodecyl sulfate-polyacrylamide gel electrophoresis and transferred to a nitrocellulose membrane. The membrane was incubated for 20 min in Tris-buffered saline containing 0.1% Tween 20 (TBS-T) supplemented with 5% skim milk f or blocking, f ollowed by overnight incubation with a primary antibod y a t 4 • C . The blots were washed and incubated with horseradish peroxidase-conjugated secondary antibody (Enzo Life Sciences) in TBS-T at 1:5,000 dilution for 1 h. Signals were detected using an enhanced chemiluminescent reagent (Thermo Fisher Scientific) by an automated imaging system (ChemiDoc ™; Bio-Rad Laboratories).

Flow cytometry
Cells were washed with phospha te-buf fered saline (PBS) and fixed with 70% ethanol in PBS overnight. Fixed cells were then washed with PBS and incubated with 0.2 mg / ml RNase A in PBS at 37 • C for 1 h. DNA was stained with 10 g / ml propidium iodide in PBS. Flow cytometry was performed on a FACSVerse ™ flow cytometer using BD FAC-Suite ™ software (BD Biosciences). Data analysis was performed using the FlowJo software.

Analysis of abnormal chromosomes
Cells were incubated with 0.2 g / ml colcemid for 3 h, and then metaphase cells were harvested by trypsinization. The cells were then swollen in 0.075 M KCl at 37 • C for 15 min and fixed with methanol:acetic acid (3:1) twice. Cells were dropped onto glass microscope slides and stained with 5% Giemsa stain. Images wer e acquir ed using a fluorescence microscope (BX53; Olympus, Tok yo , Japan). At least 20 metaphase cells were taken randomly from each condition.

Sister-chromatid e x change assay
Cells were cultured in media containing BrdU at a final concentration of 25 g / ml for 48 h. CDEAH was added 24 h before harvest, and colcemid (0.2 g / ml) was added for the final 3 h. Metaphase cells were harvested by trypsinization. The cells were then swollen in 0.075 M KCl at 37 • C for 15 min and fixed with methanol:acetic acid (3:1) twice. Cells were dropped onto glass microscope slides and stained with 5% Giemsa stain. Images were acquired using a fluorescence microscope (BX53; Olympus). At least 20 metaphase cells were taken randomly from each condition.

Apoptosis assay
Apoptotic cell death was quantified using an Annexin V Alexa Fluor ™ 488 conjugate (A13201, Thermo Fisher Scientific) and a BD FACSVerse instrument with FlowJo software (version 10) according to the manufacturer's instructions.

Comet assay
The comet assay was performed using a CometChip ® (Tre vigen) accor ding to the manufacturer's instructions. In brief, single-cell suspensions were prepared in 6 ml medium with 1.0 × 10 5 cells / ml density. Aliquots of 100 l cells per well were applied to a CometChip and incubated in a tissue culture incubator for 10 min, with gentle shaking three times in 10 min intervals to spread cells e v enly. Medium was removed and each CometChip from the 96well CometChip ® system was gently washed with 5 ml PBS twice. The CometChip was then covered with 6 ml of 1% 45 • C low-melting agarose in PBS. After the solidification of the agarose, the slide was immersed in a lysis solution (Tre vigen) ov ernight a t 4 • C . The CometChip was equilibrated twice in an alkaline solution at 4 • C for 20 min, electrophoresed at 4 • C for 50 min at 22 V in an alkaline solution, neutralized twice at 4 • C for 15 min in fresh 0.4 M Tris (pH 7.4) buffer and then equilibrated at 4 • C for 30 min in 20 mM Tris (pH 7.4) buffer. DNA in CometChips was stained with 0.2 × SYBR ® Gold in 20 mM Tris (pH 7.4) buf fer a t room tempera ture for 2 h. Images wer e acquir ed with a fluorescence microscope (BX53; Olympus) and the tail moment was calculated using the Comet analysis softwar e (Tr evigen).

Cell viability assay
Cells were plated in white, solid-bottom 96-well plates for a 2-day incubation period. HAP1 and HCT116 cells were pla ted a t a final density of 5,000 cells per well, while XP2OS cells were plated at a density of 3,000 cells per well and incubated for 1 day prior to treatment with the specified compounds. Cell viability was measured 2 days after treatment using Cell Titer-Glo (Promega) according to the manufacturer's instructions.
For cell viability assay after a 6-day incubation protocol, cells were plated in black, solid-bottom 96-well plates. HAP1 cells were plated at a final density of 600 cells per well, while TK6 cells and HCT116 cells were plated at a final density of 700 cells per well and incubated for 1 day prior  to treatment with the indicated compounds. Cell viability was measured 6 days after treatment using Cell Titer-Blue (Promega) according to the manufacturer's instructions. Viability was quantified in a Synergy NEO2 Hybrid Multi-Mode Reader (BioTek). The Chou-Talalay combination index method ( 37 ) was utilized to assess the effects of the drug combination. The analysis was performed using the freely accessible CompuSyn software tool.

Alkylation of nucleobases with CDEAH
Reaction of adenine with CDEAH. CDEAH (28.8 mg, 0.2 mmol, 1.0 equiv.) was added to a solution of adenine (27.0 mg, 0.2 mmol, 1.0 equi v.) dissolv ed in TFE (2.0 ml) (see Scheme 1 ). Sodium acetate (24.6 mg, 0.3 mmol, 1.5 equiv.) was added to adjust the pH to neutral. The r eaction mixtur e was stirr ed at 37 or 60 • C for 3 days. After incubation, the crude reactant was filtered through a syringe filter to r emove pr ecipita tes. The filtra ted chemicals were characterized by ultra-performance liquid chromato gra phy-high-resolution accurate mass-parallel reaction monitoring (UPLC-HRAM-PRM) as described below.
Reaction of guanine with CDEAH. CDEAH (28.8 mg, 0.2 mmol, 1.0 equiv.) was added to a solution of guanine (30.2 mg, 0.2 mmol, 1.0 equi v.) dissolv ed in TFE (2.0 ml) (see Scheme 2 ). Sodium acetate (24.6 mg, 0.3 mmol, 1.5 equiv.) was added to adjust the pH to neutral. The reaction mixture was stirred at 37 or 60 • C for 3 days. After incubation, the crude reactant was filtered through a syringe filter to r emove pr ecipita tes. The filtra ted chemicals were characterized by UPLC-HRAM-PRM as described below.
Reaction of thymine with CDEAH. CDEAH (28.8 mg, 0.2 mmol, 1.0 equiv.) was added to a solution of thymine (25.2 mg, 0.2 mmol, 1.0 equi v.) dissolv ed in TFE (2.0 ml) (see Scheme 3 ). Sodium acetate (24.6 mg, 0.3 mmol, 1.5 equiv.) was added to adjust the pH to neutral. The reaction mixture was stirred at 37 or 60 • C for 3 days. After incubation, the crude reactant was filtered through a syringe filter to r emove pr ecipita tes. The filtra ted chemicals were characterized by UPLC-HRAM-PRM as described below.
Reaction of cytosine with CDEAH. CDEAH (28.8 mg, 0.2 mmol, 1.0 equiv.) was added to a solution of cytosine (22.2 mg, 0.2 mmol, 1.0 equi v.) dissolv ed in TFE (2.0 ml) (see Scheme 4 ). Sodium acetate (24.6 mg, 0.3 mmol, 1.5 equiv.) was added to adjust the pH to neutral. The reaction mixture was stirred at 37 or 60 • C for 3 days. After incubation, the crude reactant was filtered through a syringe filter to r emove pr ecipita tes. The filtra ted chemicals were characterized by UPLC-HRAM-PRM as described below.

SPE purification of alkylated nucleobases
An aliquot of each alkylated N-[dieth ylamino(eth yl)]n ucleobase (DEAE-n ucleobase) r eaction was r econstituted in 1 ml of 5% methanol in water and sonicated for 30 min. Following sonication, the solution was centrifuged at 14,000 rcf at room temperature for 10 min to pellet the solid precipitate. Oasis ® HLB 30 mg extraction cartridges (Waters, Milford, MA) were placed on a vacuum manifold and conditioned with two additions of 1 ml of water, and then 1 ml of methanol with a gentle vacuum applied. The sample solutions were then loaded onto a column, followed by washing twice with 2 ml of 5% methanol in water. DEAE-purine nucleobases and DEAE-pyrimidine nucleobases were eluted with two additions of 500 l of 100% methanol. Collected elution was concentrated by centrifugal vacuum and stored at −20 • C for future analysis.

Alkylation of calf thymus DNA with CDEAH
An aliquot of 100 g of calf thymus DNA (CTDNA) dissolved in water or PBS was diluted to 190 l with water or PBS, followed by adding 10 l of 2 mM CDEAH to yield a final concentration of 100 M. The solutions were then incuba ted a t 37 • C for 16 h to allow alkyla tion of nucleotides, followed by heating at 70 • C for 1 h to depurinate DEAEpurine bases. The released DEAE-purine nucleobases were separated from the DNA backbone by centrifugation at 14,000 rcf at 4 • C for 10 min through Nanosep ® centrifugal devices with Omega ™ 10-kDa membranes. The filters were further washed using an equal volume of deionized (DI) water twice, and 100 l of 50:50 acetonitrile (ACN):DI water once. All collected solutions (depurination solution) were concentrated to dryness by centrifugal vacuum and stored at −20 • C for future experiments. The DNA backbone in the filter was resuspended in 100 l water (DNA backbone solution), r ecover ed from the filter and stored at −20 • C for future experiments.

Characterization of alkyl-nucleobase standards by UPLC-HRAM-PRM
Alkylation of purine and pyrimidine nucleobases by CDEAH was confirmed by analyzing the solid-phase extraction (SPE)-purified reaction products by a UPLC-HRAM-PRM assay in positi v e mode as follows: a Hypersil GOLD 1.9 m C18 column (100 mm × 1.0 mm) was operated using a gradient of buffer A (15 mM ammonium acetate, pH 7.0) and buffer B (100% acetonitrile) at 0.05 ml / min starting at 2% buffer B for 2 min, linearly increased to 25% buffer B over 8 min, followed by an increase to 50% buffer B over 20 min, then an increase to 80% buffer B over 2 min, held constant at 80% buffer B f or 2 min, f ollowed by a decrease to 2% buffer B over 2 min and finally re-equilibrated at 2% buffer B for 9 min. Mass spectrometry (MS) settings were as follows: electrospray voltage, 3,500 V; capillary temperature, 320 • C; full scan AGC, 1 × 10 6 ; full scan resolution, 70,000; HESI tempera ture, 150 • C; shea th gas, auxiliary gas and sweep gas flow rates, 35

Analysis of CTDNA alkylation by CDEAH by UPLC-HRAM-PRM
The depurination solution from above was reconstituted in 50 l water and measured by a microvolume UV spectrophotometer (Thermo Scientific ™ NanoDrop) using the extinction coefficient for guanosine to confirm that nucleobases were present. The equivalent of 1.2 g of CTDNA from the depurination solution was analyzed by the DEAEpurine UPLC-HRAM-PRM method described above.
The DNA backbone solution from above was measured by a microvolume UV spectrophotometer, and a 25 g aliquot of CTDNA was diluted to 150 l of 1 × NEB nucleoside digestion mix reaction buffer and incubated with 2.5 l NEB nucleoside digestion mix (1 l mix per 10 g CTDNA) at 37 • C for 4 h. Following incubation, the digestion enzymes were removed by centrifugation at 14,000 rcf at 4 • C for 10 min through Nanosep ® centrifugal devices with Omega ™ 10-kDa membranes. The filters were further washed using an equal volume of DI water one Tumor sizes wer e measur ed using calipers e v ery 3 days for 16 days following drug treatment. Tumor volume was measured by the following formula: V = ( width ) 2 × length × (1 / 2) . All mice were euthanized to harvest tumors for immunostaining.

Immunohistochemistry
Hematoxylin and eosin (H&E) staining, TUNEL and ␥ -H2AX immunostaining were commercially performed by Histoire (Seoul, Republic of Korea). Collected tumors were fixed in formalin and picked up by Histoire. Detailed immunostaining procedures can be checked by accessing the Histoire's w e bsite ( http://www.histoire.co.kr/ ).

CDEAH selectively kills XPA-and PARP1-deficient cells
We previously developed a high-throughput genotoxicity screening assay that uses ATAD5 expression as a biomarker to identify genotoxic compounds ( 33 ). Using this assay, we identified CDEAH (Figure 1 A) that increased the ATAD5luciferase expression (Figure 1 (Figure 1 D). We also measured cell viability in XP2OS cells deri v ed from an XPA mutant xeroderma pigmentosum patient and found that they showed significant and dose-dependent sensitivity to CDEAH (Figure 1 D). Sensitivity of XP2OS cells to CDEAH was rescued by expression of WT XPA protein complemented with the WT XPA, and the combination of CDEAH and olaparib induced an additi v e ef fect in XPA KO ra ther than in WT (Supplementary Figure S2A Figure S1D).
To investigate whether NER is necessary to repair the CDEAH-induced lesions, we conducted a cell viability assay of CDEAH in XPC KO and CSB KO cell lines. We found that CDEAH induced mild sensitivity in XPC KO cells, but se v ere sensiti vity in CSB KO cells (Figure 1 G), suggesting that CDEAH induces lesions that are dependent more on transcription-coupled NER compared to the global genome NER.
We also tested the effect of CDEAH on lowering the dosage of olaparib to specifically kill BRCA1 / 2 -deficient cells by performing cell viability assay using TK6 cells with BRCA2-mAID-GFP. We found that incubating this cell with auxin, which degrades BRCA2 protein, resulted in sensitivity to olaparib in a dose-dependent manner. Moreover, co-treatment of CDEAH and olaparib induced hypersensitivity specifically in BRCA2 -deficient cells (Figure 1 H).
Taken together, the result suggests that CDEAH can selecti v ely cause hypersensiti vity in cells defecti v e in NER and PARP1-dependent BER pathways.

Characterization of the reaction of CDEAH with nucleobases
CDEAH is half-nitrogen mustard, and its Cl group can be displaced by an intramolecular ring-closing reaction to yield a highly electrophilic aziridinium ion, which can alkylate the nucleophilic positions of DNA nucleobases. To test this potential mechanism, we investigated the alkylation of purine and pyrimidine nucleobases by CDEAH by in-

Confirmation and characterization of alkylated CTDNA by CDEAH
To further investiga te CDEAH alkyla tion on nucleotides in double-stranded DNA, CTDNA was incubated with CDEAH for 16 h. Alkylated purines wer e r eleased from the DNA backbone by thermal hydrolysis and analyzed by the UPLC-HRAM-PRM assays ( 39 , 40 ). When the equivalent of 240 ng of CTDNA was analyzed, we were able to confirm the presence of DEAE-guanine at 10.3 min (Figure 2 B). Since the N9 position of guanine is not accessible in double-stranded DNA, the observed peak at 10.3 min is expected to be N7-DEAE-guanine and the standard peak at 12.1 min is expected to be N9-DEAE-guanine. During the analysis of 240 ng of CTDNA, we were unable to detect any DEAE-adenine. When the scale of the reaction was increased 5-fold to 1.2 g of CTDNA, we were able to detect a weak signal for DEAE-adenine at both 13.1 and 14.3 min (Figure 2 B). Analysis of the PRM data re v ealed that both peaks had the expected fragmentation pattern. Gi v en that the N3 position of 2 -deoxyadenosine is the most reacti v e position with other nitro gen m ustards in DNA ( 38 , 41 ), we belie v e the weak signal at 13.1 min corresponds to the N7-DEAE-adenine and the stronger signal at 14.3 min corresponds to N3-DEAE-adenine.
The remaining alkylated DNA backbone was digested with NEB nucleoside digestion mix to yield DEAE-2deoxypyrimidines. When 5 g of CTDNA digestion was analyzed by the modified DEAE-pyrimidine nucleoside UPLC-HRAM-PRM assay, w e w ere able to detect two DEAE-dC peaks at 11.9 and 13.3 min and one DEAE-dT  Supplementary Figure S3B). Gi v en that the N1 position of pyrimidines is not accessible in nucleotides, the observed peak at 11.9 min is expected to correspond to O 2 -DEAE-dC and the peak at 12.1 min is expected to correspond to O 2 -DEAE-dT or O 4 -DEAE-dT. Taken together, CDEAH is most reacti v e to 2deo xyguanosine, follo wed by 2 -deo xyadenosine and then 2 -deoxypyrimidines ( dG dA > dC ∼ dT ). To investigate reactivity and alkylation potential, we treated WT and mutant HAP1 cell lines with a CDEAH deri vati v e that replaced the chloride-leaving group with a br omide-leaving gr oup (Supplementary Figure S1A and B). Howe v er, there was no significant increase in sensitivity with the bromide-substituted deri vati v e compared to CDEAH.

CDEAH induces more DNA breaks in PARP1-deficient cells
Since CDEAH alkylates nucleobases, CDEAH treatment is expected to interfere with replication and S-phase progression. We investigated the effect of CDEAH on cell cycle progression in HCT116 WT and PARP1 KO cell lines. Both WT and PARP1 KO cells were arrested at S phase upon treatment with increasing concentrations of CDEAH, but there was no significant difference in WT and PARP1 KO cells (Figure 3 A). We confirmed that CDEAH also increases PAR, as previous studies have reported ( 42 , 43 ) an increase in PAR le v els by alkylating agents (Supplementary Figure S5A). Since PARP1 KO cells were selecti v ely killed by CDEAH, we compared DNA damage markers after 24h treatment of 80 M CDEAH in WT and PARP1 KO cells. Consistent with cell viability results, we found higher ␥ -H2AX induction in CDEAH-treated PARP1 KO cells (Figur e 3 B). Furthermor e, actual single-stranded DNA br eaks measured by the alkaline comet assay wer e incr eased in PARP1 KO cells compared to WT cells upon CDEAH tr eatment (Figur e 3 C). To investigate genomic instability, we tested sister-chromatid exchange (SCE) frequency and abnormal chromosomes in HCT116 WT and PARP1 KO cells after 24-h treatment with 80 M CDEAH. Consistent with a previous study, PARP1 -deficient cells showed incr eased SCE fr equency e v en in the absence of exogenous damage (44)(45)(46)(47). We found a further increase in SCE frequency and abnormal chromosomes in CDEAH-treated PARP1 KO cells (Figure 3 D-F). For example, cells with > 25 breaks per metaphase were significantly increased in CDEAH-treated PARP1 KO cells (Figure 3 G). Finally, apoptotic cell death was quantified using an Annexin V Alexa Fluor ™ 488 conjugate. Compared to the WT, the PARP1 KO cell line showed significantly increased apoptosis upon treatment with 80 M CDEAH (Figure 3 H). These results show that DNA damage, chromosomal aberrations and apoptotic cell death are increased in PARP1 -deficient cells following treatment with CDEAH.

Growth of PARP1 KO xenograft tumors is selectively inhibited by CDEAH treatment
To determine the effect of CDEAH on tumor growth in vivo , we analyzed the susceptibility of tumor xenografts in nude mice to treatment with CDEAH (Figure 4 A)

DISCUSSION
In the present study, we identified CDEAH as an agent that selecti v ely kills PARP1-and XPA-deficient cells in vitro and in vivo . Our data suggest that co-treatment of PARPi such as olaparib with CDEAH enhances tumor growth inhibition. CDEAH pr efer entially alkylates guanine DNA bases, w hich subsequentl y interferes with pro gression through S phase, which is further enhanced in PARP1-deficient cells leading to cell death. The PARP1 dependence of CDEAH covalent adducts gi v es a potential synergistic combination treatment option with P ARPi. P ARPi such as olaparib, ruca parib, nira parib, talazoparib and veliparib are FDA approved for clinical usage ( 17 ). In addition to monotherapy of these PARPi, various combinatorial treatments with PARPi ar e curr ently being investigated in clinical trials. For example, TMZ has been used in combinatorial therapy with rucaparib e v er since the first clinical trial of a PARPi in 2003 ( 17 ). Increased sensitivity of cells with reduced PARP1 activity to the TOP1 inhibitor camptothecin, irinotecan and topotecan, which is used as a cancer therapeutic agent, raised a potential combinatorial treatment of TOP1i with PARPi ( 11 , 17 , 48-52 ). In addition, various combinatorial therapies are currently undergoing clinical trials for PARPi in various cancers ( 11 , 17 ).
Ther e ar e many other well-characterized DNA alkylating agents, such as nitrogen mustard compounds, cisplatin and MMS (53)(54)(55)(56)(57). Nitrogen mustard compounds and cisplatin have two electrophiles (Cl ligands) and form an interor intr astr and cross-link between two nucleobases in addition to monoadducts. Such complex adducts induce various higher toxic effects on both tumor and normal cells  that need to be overcome by either the NER or inter-crosslink repair pathwa y f or cell survival (55)(56)(57)(58)(59)(60)(61). In contrast to these agents, since CDEAH possesses a single electrophile, it makes only one type of DNA ad duct, monoad duct. These simple adducts are substrates for either the BER or NER pathwa ys, and thus ma y induce milder side effects on normal cells. The alkylating agent MMS makes a simple monoadduct with DNA nucleobases similar to CDEAH ( 53 , 54 ). Pre vious e xperiments reported a synergistic effect of 0.01% MMS with olaparib ( 62 , 63 ), whereas XPA deficiency did not show an effect ( 64 , 65 ). To confirm these, we incubated HAP1 cell lines with PARP1 or XPA deficiency with MMS or TMZ for 48 h and measured cell viability. Our results showed that only the PARP1 KO HAP1 cell lines exhibited significant sensitivity to MMS, while both PARP1 KO and XPA KO HAP1 cell lines had minor effects with TMZ (Supplementary Figure S4A and B). In contrast, the CDEAH sho ws syner gistic effect with olaparib and selecti v ely kills not only PARP1-but also XPA-deficient cells (Figur e 1 ). A pr evious study r eported that monoadducts of melphalan with two electrophiles (Cl ligands) can be repaired by NER in human cell-free extracts ( 66 ). Taken together, CDEAH covalent adducts are bulky and subject to NER. Lastly, the high water solubility of CDEAH compared to other cross-linking reagents is another beneficial characteristic of CDEAH for combinatorial treatment.
CDEAH sensitizes cells defecti v e in either NER or PARP1-dependent BER pathways. Thus, in addition to the potential clinical uses of CDEAH for cancer therapy, CDEAH can be used as a tool compound to better understand NER and PARP1-dependent BER pathways in detail. W hen CDEAH alkyla tes the DNA nucleobase, a very strong covalent bond is formed between CDEAH and the nucleobase, especially guanine. CDEAH adducts should be removed from the genome by the NER or PARP1dependent BER pathway. In combination with the UPLC-HRAM-PRM assay used in this study, the removal of CDEAH adducts from the genome can be analyzed to study the kinetics of the NER and PARP1-dependent BER pathways in various DNA repair-deficient cell lines as well as cancer cells. Since remaining CDEAH adducts would cause mutations in the genome, CDEAH could also be used to study mutagenesis mechanisms in cells. CDEAH pr efer entially alkylates guanine bases, suggesting that a unique muta tion signa ture would be produced in cells defecti v e in a differ ent DNA r epair pathway. The anal ysis of m utation signa tures accumula ted in CDEAH-trea ted cancer cells with different genetic backgrounds could be used as r efer ences to choose treatment options.
CDEAH tr eatment incr eased the population of S-phase cells. This could be due to the stalling of DNA replication by the CDEAH-alkylated nucleobases. DNA damage in the genome at the S phase is recognized by the MMR machinery or bypassed by the translesion synthesis (TLS) pathway ( 67 ). It would be interesting to investigate whether CDEAH adducts at guanine induce effects on the MMR pathway. CDEAH adducts encountering DNA replication forks would be bypassed by the TLS pathway. Since TLS is an err or-pr one pa thway tha t genera tes muta tions in the genome, it will be interesting to investigate what type of mutations would be produced by CDEAH and which TLS enzymes are involved in mutagenesis. In addition to DNA replication, CDEAH adducts in the coding sequences could affect transcription. The pr efer ential alkylation of guanine bases suggests that CDEAH would target CpG islands as a transcriptional obstacle. Most CpG islands in promoter are unmethylated, but in the silenced promoter, CpG islands are usually methylated and are important for the silencing state of the promoter (68)(69)(70). CDEAH could impact this regulation as well. It will be interesting to investigate the transcriptional impact of CDEAH on genes carrying CpG island in their promoters. Other sequence-dependent DNA structures in the genome, such as G-quadruplex-rich sequences frequently found in telomeres ( 71 , 72 ), could be affected by CDEAH as well.
Collecti v ely, we identified a small molecule, CDEAH, as a potential sensitizing agent for PARPi and a tool compound to study various DNA repair pathways. The de v elopment of sensitizers using the synthetic lethality strategy used in this study suggests that many other small molecules can be de v eloped to target various genetic diseases, including cancers.

DA T A A V AILABILITY
The data underlying this article are available in the article and in its online supplementary material.

SUPPLEMENT ARY DA T A
Supplementary Data are available at NAR Cancer Online.